Genetic Knock-Ins of Endogenous Fluorescent Tags in RAW 264.7 Murine Macrophages Using CRISPR/Cas9 Genome Editing

CRISPR/Cas9 genome editing is a widely used tool for creating genetic knock-ins, which allow for endogenous tagging of genes. This is in contrast with random insertion using viral vectors, where expression of the inserted transgene changes the total copy number of a gene in a cell and does not reflect the endogenous chromatin environment or any trans-acting regulation experienced at a locus. There are very few protocols for endogenous fluorescent tagging in macrophages. Here, we describe a protocol to design and test CRISPR guide RNAs and donor plasmids, to transfect them into RAW 264.7 mouse macrophage-like cells using the Neon transfection system and to grow up clonal populations of cells containing the endogenous knock-in at various loci. We have used this protocol to create endogenous fluorescent knock-ins in at least six loci, including both endogenously tagging genes and inserting transgenes in the Rosa26 and Tigre safe harbor loci. This protocol uses circular plasmid DNA as the donor template and delivers the sgRNA and Cas9 as an all-in-one expression plasmid. We designed this protocol for fluorescent protein knock-ins; it is best used when positive clones can be identified by fluorescence. However, it may be possible to adapt the protocol for non-fluorescent knock-ins. This protocol allows for the fairly straightforward creation of clonal populations of macrophages with tags at the endogenous loci of genes. We also describe how to set up imaging experiments in 24-well plates to track fluorescence in the edited cells over time. Key features • CRISPR knock-in of fluorescent proteins in RAW 264.7 mouse macrophages at diverse genomic loci. • This protocol is optimized for the use of the Neon transfection system. • Includes instructions for growing up edited clonal populations from single cells with one single-cell sorting step and efficient growth in conditioned media after cell sorting. • Designed for knocking in fluorescent proteins and screening transfected cells by FACS, but modification for non-fluorescent knock-ins may be possible.


Recipes
To make the 0.1 M EDTA, dissolve the EDTA powder in 45 mL of autoclaved MilliQ water, then top off to 50 mL once totally dissolved for a total volume of 50 mL.Filter this solution through a 0.2 μm syringe filter (Acrodisc) before use.

FACS sorting buffer
Reagent Final concentration Quantity or Volume DPBS n/a 47.75 mL EDTA 0.1 M (from Recipe 4) 1 mM 500 µL HEPES 1 M 25 mM 1.25 mL FBS 1 % 500 µL Total n/a 50 mL To make the FACS sorting buffer, combine all reagents and then pass through a 0.2 μm syringe filter (Acrodisc).Note: Other compositions of FACS sorting buffer will likely also work.n/a 10 mL To make the 1,000× puromycin, dissolve the puromycin dihydrochloride in 9 mL of autoclaved MilliQ water, then top off to 10 mL once totally dissolved for a total volume of 10 mL.Filter this solution through a 0.2 μm syringe filter (Acrodisc) before use and store at -20 °C in 1 mL aliquots.

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Published: Mar 20, 2024 d.Select guides with high specificity scores and as high as possible cutting efficiency scores.You should additionally optimize for being as close to your desired insertion site as possible.You should select the three most promising guides (Figure 2).Note: Rarely do you have guides that look perfect here.This is part of why we screen three guides, so that you can choose several that look promising and then test them.The cutting efficiency prediction algorithms are not accurate for all cell types or loci, and this is also why we test the guides.e. Order two oligos (one forward and one reverse) per guide (from Eurofins, IDT, etc.).For the forward oligo, append CAACG to the 5' end of the guide sequence.For the reverse oligo, take the reversecomplement of the forward guide sequence (not including the added CAACG) and add AAAC to the 5' end and C to the 3' end.For example, if your guide sequence was ACCCTT…GGGCT, then you would order forward oligo caacgACCCTT….GGGCT and reverse oligo aaacAGCCC…AAGGGTc.Note: Other types of competent cells will likely also work.d.Pick three colonies from the plate that has grown up overnight and grow up each picked colony in a 5 mL culture of LB + carbenicillin.Perform minipreps using the QIAprep Spin Miniprep kit to extract the plasmids from the bacteria and sequence the plasmids using a universal U6 primer to confirm correct gRNA insertion.Re-streak the minipreps with the correct plasmid sequence onto new LBcarbenicillin plates and grow up overnight.e. Pick a colony from the re-streak plate that had the correct sequencing and grow up this colony in a midiprep culture of 100 mL of LB + carbenicillin (so adding 100 μL of 1,000× carbenicillin).The culture should be grown overnight at 37 °C with shaking at 300 rpm.f.Use the Macherey Nagel NucleoBond Midi Plus EF kit to perform a midiprep to extract the plasmid from the midiprep culture.Follow the kit instructions for midipreps of low-copy plasmids.For clarification and loading, we load the entire mix onto the filter (rather than spinning it out first); for DNA concentration at the end, we use the finalizers.We elute in 600 µL of provided water.This resulting plasmid is the pSp-Cas9-puro gRNA plasmid that you will use in subsequent steps.Note: Other midiprep kits could likely be used, but it is important for the final DNA concentration to be high, ideally over 1 μg/μL.For transfection later into macrophages, it is also important for the DNA to be endotoxin-free, which this kit accomplishes.

Test gRNA cutting efficiency
a. On day 1, use your hemocytometer to count the cells and seed 2 × 10 5 NIH3T3 cells per well in a 6well plate in complete DMEM.Seed the same number of wells as gRNAs that you are testing (usually three) plus one well as a control for uncut genomic DNA and one well as a control for puromycin selection.
Note: We use NIH3T3 cells to test the gRNA cutting efficiency because they can be transiently transfected with a high efficiency and therefore selected for transfected cells while the cells are still expressing the transient selectable marker.RAW 264.7 cells transfected using the Neon system are too sick immediately after transfection to be selected using puromycin.We find that gRNAs selected for high efficiency in NIH3T3 cells are very effective for creating knock-ins in RAW 264.7 cells.Note: Be gentle with this solution, as the DNA-lipofectamine complex can be fragile.iv.Add the transfection solution dropwise onto the well, gently tilt the plate to mix, and return the plate to the incubator.c.On day 3, trypsinize the wells and transfer the entire contents of each well into its own 10 cm plate in complete DMEM.For the gRNA test plates and the puromycin control plate, add 10 μL of 1 mg/mL puromycin in 10 mL of total media.d.On day 5, inspect the puromycin control plate and confirm that all cells are dead.Wash the gRNA test plates with DPBS and add fresh complete DMEM (without puromycin) onto the plate.Split the uncut genomic DNA control plate to be confluent the next day (usually 1:3).e.On day 6, collect the cells using trypsin (each condition of gRNA test cells and the uncut control), spin down into cell pellets, and store at -80 °C.Note: Occasionally, these primers will be bad sequencing primers; in this case, you can design and order an additional internal primer to sequence across the cut site.f.Use the online Synthego ICE tool (https://ice.synthego.com/#/)to assess gRNA cutting efficiency.
Note: This tool compares the .ab1Sanger sequencing files in uncut (control) cells and cells transfected with a gRNA to determine the fraction of cells with an indel at the CRISPR site.The software also requires you to input the guide sequence used for that sample.The fraction of cells with an indel correlates with the gRNA cutting efficiency, which is also calculated by the software.The software also shows exactly which indels were formed (Figure 3).g.Choose the gRNA with the best cutting efficiency that is as close to your desired insertion site as possible.
Note: We select the gRNA with the highest cutting efficiency (indel percentage) that is as close to our desired insertion site as possible.In our experience, any indel percentage above 50% is generally sufficient, though we have not specifically tested moving forward with a gRNA with a score of 40% vs. 60%, for example.We have seen gRNAs with an indel score below 30% fail, though we cannot  For N-terminal tagging, the insertion site should be immediately after the endogenous ATG start codon and, for C-terminal tagging, it should be immediately before the endogenous stop codon.Once the best gRNA has been identified, its PAM sequence needs to be mutated in the donor plasmid to avoid re-cutting.We do this using primers with overhangs to add new sequence.If the PAM cannot be mutated synonymously, then at least three synonymous mutations should be made in the seed sequence of the gRNA so that Cas9 re-binding is prevented in that way.If the PAM is in a UTR, we do a literature search to identify if it is in a region of the UTR known to be important for regulation; if not, we simply mutate the PAM.If there is known regulation in the UTR, we try to avoid mutating that region.The desired fluorescent protein insertion sequence also needs to be amplified via PCR from a plasmid containing that fluorescent protein.We assemble the plasmids using Gibson assembly in a pUC19 backbone and so add the appropriate Gibson overhangs on our primers as well when needed.This generally results in a four-part Gibson assembly reaction with the plasmid backbone, left homology arm, right homology arm, and fluorescent protein (Figure 4).  the fluorescent tag, these can also be introduced on primer tails when designing your PCR.We use the T2A sequence from Nora et al. [6] and a GDGAGLIN linker with DNA sequence GGCGACGGCGCCGGCCTGATCAAC. 2. Transform the cloned plasmid (after Gibson assembly) into NEB 5-alpha competent E. coli (high efficiency) as above.3. Grow up four minipreps, sequence the minipreps, and identify a miniprep with the correct sequencing.
You want to be sure the entire homology arm and insertion sequences are correct.4. Grow up a midiprep culture of the plasmid with the correct sequencing and perform the midiprep using the Macherey Nagel NucleoBond Midi Plus EF kit in the same way as described for the gRNA-Cas9 plasmid above.Note: Plasmid concentration is best if above 1 μg/μL.If it is below 500 ng/μL, we suggest repeating the midiprep.
C. Transfect gRNA-Cas9 plasmid and donor plasmid into RAW 264.7 cells (Figure 5) Note: We use the Neon transfection system rather than any form of lipofection because, in our hands, the Neon transfection system gives much higher transfection efficiency than any lipofection method in RAW 264.7 cells.See General Note 6.
1. Have a confluent 10 cm plate of RAW 264.7 cells ready on the day of transfection.
2. Fill three wells of a 6-well plate with 2 mL each of antibiotic-free DMEM.Note: It is important to use the antibiotic-free DMEM here, as having antibiotics in the media decreases cell viability after transfection.3.In an Eppendorf tube, combine 45 μg of gRNA-Cas9 plasmid and 45 μg of donor plasmid.Ideally, both the donor and gRNA-Cas9 plasmid concentrations are over 1 μg/μL.a.In this protocol, we transfect in triplicate, so each single transfection is 15 μg of each plasmid.

Note: We attempted various ratios of gRNA-Cas9 DNA to donor DNA, different volumes of DNA, pretreating the cells with DMSO, pretreating with interferon-gamma (as our genes were interferongamma inducible and the idea was to open up the chromatin at their loci), and also plating posttransfection into media containing 2 μM M3814 (M3814 is a DNA-PK inhibitor that can increase the
Published: Mar 20, 2024 ratio of HDR edits to NHEJ edits [13]).Adding more DNA decreased the cell viability after transfection, and we saw little effect of either DMSO pretreatment or M3814 post-treatment.Therefore, we chose not to either pretreat or post-treat our cells for our experiments.Note: We do not linearize our plasmids before transfection, but we did not test how linearization might affect transfection or knock-in efficiency.One reason we did not attempt plasmid linearization was concern over the possible introduction of endotoxins in this process.Therefore, we are transfecting circular plasmid DNA.a.In this protocol, we transfect in triplicate, so each single transfection is 2.5 × 10 6 cells.7. Resuspend cells in 1 mL of DPBS, spin down again, and aspirate the supernatant.8. Resuspend the cells in 300 μL of R buffer from the Neon electroporation reagent kit.
Note: The kit says that extended R buffer exposure can be harmful for cells, so move quickly after this step.9. Add the cells in R buffer to the tube with your plasmid DNA and pipette up and down gently 3-5 times.10.Use a 100 μL tip on the Neon pipette to pick up 100 μL of the cells-buffer-DNA solution.Make sure there are no bubbles in the tip.11.Put the tip in the Neon cuvette and push down until it clicks.12. Press Start on the program (1,680 V, 20 ms, 1 pulse).13.Once the program has completed, gently eject the contents of the tip into one well of a 6-well plate with antibiotic-free DMEM.14. Insert the pipette tip into the tube of DPBS and pipette the DPBS up and down five times to wash.15.Repeat steps C10-14 twice more, for a total of three transfections, plating into a new well for each transfection.16.Once all transfections are complete, rock the cell plate back and forth to evenly distribute cells.17.Place the cell plate in your tissue culture incubator.18.Put away the Neon machine.According to Thermo Fisher, the tips are not reusable, and the cuvettes should only be reused a few times.In our experience, tips can be reused at least five times, but we use a new tip for each new cell line that we construct to avoid contamination.To store tips, we place them in a 15 mL conical tube.At least 24 h before reuse, we use forceps to separate the internal gold plunger from the external plastic, put both the plunger and plastic back in the 15 mL conical tube, and add filtered 100% ethanol for at least 2 h.We then dry them by placing them on a Kimwipe in a tissue culture hood with the UV light on for 1 h.We also reuse cuvettes at least five times, though we do sometimes see a decrease in 13 Published: Mar 20, 2024 cell viability after transfection using older cuvettes.We store cuvettes in a 50 mL conical tube with filtered 100% ethanol, and before reuse we remove them from the conical tube and dry them by placing them on a Kimwipe in a tissue culture hood with the UV light on for 1 h.

D. Grow up cells, sort single positive cells, and grow up clonal colonies from single cells
1. Two days after electroporation, inspect cell viability.There may be many floating cells, many of which are dead but some of which may be alive.Note: Cell viability after transfection can vary greatly based on a number of factors.It can depend on cell health before electroporation, age of reagents (buffers, cuvettes, etc.), as well as the nature and purity of the DNA.We have done electroporations of the same DNA into two different RAW 264.7 lines in parallel on the same day and had very different post-transfection viability between them.We have also electroporated different DNAs into the same cell line on the same day and had different viability.At two days post-transfection, an electroporation that had good viability will look like a ~50% confluent well of cells, but the cells are more elongated than they normally are, so 50% confluence here is fewer cells than 50% confluence of normally growing RAW 264.7 cells.An electroporation that had poor viability will look like sparse single cells on the bottom.However, we have still seen successful genome editing in cases of poor viability after transfection.2. Use a cell scraper followed by strongly pipetting media against the bottom of the well to collect all cells in all three wells.Combine these into one 15 mL conical tube, spin down the conical tube at 200× g for 3 min, and aspirate the supernatant.3. Resuspend the cells in 2 mL of complete DMEM and place this suspension in a new well of a 6-well plate (so you have now combined three wells into one well).4. Monitor the well until it reaches confluence; this generally takes 2-5 days depending on cell viability.5. Expand these cells into a confluent 10 cm plate.6.We like to first inspect the cells under a microscope to see if they express the knocked-in fluorescent protein before sorting them.To do this, we plate two wells of a 6-well plate with these bulk transfected cells and image 60 positions per well every hour for 24 h, adding our stimulus after 2 h for stimulusinducible genes.If you see the fluorescence expression, that is great!If not, sometimes the actual knockins are so rare that you can only pick up on them via FACS; we have had cases where we do not see fluorescence via microscopy but do pick up correctly edited cells via FACS.Be sure to only image a fraction of your bulk transfected population and keep most of it in the incubator to prepare for sorting (Figure 6).7. Sort single positive cells into a 96-well plate.
Note: We generally sort 7-9 days after transfection.Occasionally, it will take longer for the cells to grow up sufficiently; that is ok too.a. Induce a 10 cm plate of transfected cells with whatever stimulus should express your fluorescent knock-in.If it is constitutively expressed, then skip this step.b.Prepare two 96-well plates with 100 μL of conditioned DMEM per well.
Note: Using conditioned rather than regular DMEM results in the single cells growing into colonies faster after sorting as well as a higher fraction of sorted single cells surviving and growing into colonies.This is a key step in this protocol.c.Collect your induced transfected cells, as well as the cells you made them from as a control for FACS gating.d.Spin cells down at 200× g for 3 min, resuspend in ~3 mL of FACS sorting buffer, and pass through a filter into a FACS tube (Falcon 5 mL round bottom polystyrene test tube, with cell strainer snap cap).e. Bring the FACS tubes with the cells on ice to a FACS machine.Gate for positive cells compared with the pre-transfection control.It is ok to get some false positives here because you will screen the clones further in subsequent steps.Sort cells that express your fluorescent protein higher than the control into the 96-well plates with one cell per well (Figure 7).Note: The fraction of positive cells is often very small, generally between 2% and 0.01% of the sample (see below for more discussion of this).However, this fraction is often enough to recover the knock-  In this field of view, there is a group of cells that express YFP after stimulus, which are the edited cells we are looking for.There are also many cells that do not express YFP.
Figure 7. Sample FACS plots when sorting for positive cells.In this example, iRFP+ parent cells were transfected to knock in a YFP tag.These plots show YFP vs. iRFP for each cell being sorted.On the left are the parent cells, and on the right are the bulk transfected cells.P5 is the gate used to collect positive cells that have higher YFP than any cells in the parent population.In this example, in the bulk transfected cells, P5 is 0.3% of the alive, single-cell population and 0.2% of the total population.The true YFP-positive population likely extends to the left of the P5 gate shown here, but we drew the gate here to try to minimize false positives.In other situations, it may be worth it to take some false positives to avoid throwing out true positives and to screen out the false positives later when the clones are screened by microscopy.screen by colony size and take the colonies on the bigger side.Choose 12 colonies to grow up (Figure 8).Note: Generally, if the editing worked, most (~70%-100%) clones that you screen will be positive for the fluorescence.If the editing did not work and you ended up sorting false positives, then no clones will be positive, or those that are will be positive even without induction for inducible genes.e.When the colony gets large in the 96-well plate (generally 10-12 days after sorting; it is ok to wait until the media turns yellow, but if you choose to wait that long, you should move the colony promptly after that), use forceful pipetting of media using a P200 pipette to dislodge the colony and move the cell suspension to a 24-well plate.f.Continue to grow up and transfer the cells when confluent from a 24-well plate to a 6-well plate to a 10 cm plate.

E. Screen clones
1.For the 12 colonies that you grew up, spin down two cell pellets from a confluent 10 cm plate and store at -80 °C.Additionally, freeze cryovials of cells so that you can recover ones you want to use later.2. Extract the genomic DNA from the cell pellet for each clone; we use the Zymo Quick-gDNA Miniprep kit. 3. Use PCR to amplify the region across the insertion site from the left into the right homology arm.We use the primers that we designed for determining gRNA cutting efficiency above.Use a long enough elongation time to elongate across the inserted DNA as well.We also run a control PCR on unedited wildtype DNA. 4. Run this PCR product on a gel.Note the bands: if there are two bands, then the clone is likely heterozygous for the knock-in; if there is only one large band, it is likely homozygous for the knock-in; if there is only one small band, it likely does not have the knock-in (Figure 9).

5.
Gel extract each band of DNA using the QIAquick gel extraction kit and send each band for sequencing to sequence the entire amplicon.We sequence with the same primers we used for the PCR.6. Analyze this sequencing data.For small bands, the goal is that the DNA sequence is identical to the uncut genomic DNA, rather than having been cut and repaired using non-homologous end joining to result in an indel.For large bands, the goal is to see precisely the endogenous sequence with your fluorescent protein inserted as you designed.In homozygous knock-ins, if the two alleles are slightly different, this will sometimes result in an .ab1sequencing file with overlapping peaks as there are two sequences there.In this case, sequencing from the other side or an internal primer can help.It is also the case that you want any homozygous knock-ins to be identical, and so clones with overlapping-peak sequencing may not be the best choice and you may be able to discard them (Figure 10).

F. Set up an imaging experiment
1.The day before (ideally, ~20 h before) you wish to image, seed 4 × 10 4 of your edited RAW 264.7 cells into each well of a 24-well plate in complete DMEM.We do this by preparing a tube with 104 × 10 4 cells in 13 mL of media, mixing that tube well, and then adding 500 μL of that mix to each well.After adding the cells to all wells, shake the plate back and forth on the floor of the TC hood enough to really agitate the liquid but not enough for it to spill over between wells.This is important to ensure an even distribution of cells.Incubate the plate in a tissue culture incubator for ~20 h for the cells to settle on the plate.Note: This seeding density is dense enough for the cells to be happy, but relatively sparse so that the cells can sit for 20 h in the incubator and then be imaged for 48 h without starting to grow on top of each other.You may want to modify this density based on your imaging time.2. Right before imaging, aspirate the media in each well and replace it with 500 μL of Phenol Red-free DMEM for imaging.3. Bring the plate to the microscope and set up on the microscope, including the temperature and CO2 incubation.4. Select exposure time and frequency; we take phase and iRFP images every 10 min for tracking and fluorescence in other channels every sixth cycle (every 60 min) to minimize phototoxicity. 5. Select positions, ensuring that you are in the middle of each well (which should look like the part with the darkest background).We take two images per well for a total of 48 images; our microscope cannot take more images per well within a 10 min image frequency.We also move through the plate in an "S" pattern to avoid the stage moving the plate back and forth more than necessary.

Troubleshooting
The main issue that occurs with this protocol is that you do the whole procedure and have no positive clones.In this situation, it is best to go back and make sure each previous step is working.Below are some of the intermediate steps that need to work to obtain the final knock-in.However, as discussed above, there are a couple of loci that we have been unable to tag using this protocol even after troubleshooting. Step

Neon transfection
Confirm transfection efficiency using a plasmid that constitutively expresses a fluorescent protein.
Adjust quantity of cells or DNA, ensure cells are happy before transfection, minimize time in R buffer, minimize time cells are in a tube rather than a plate, try a new Neon tip or cuvette, replace Neon buffers.

HDR at the locus
There is no direct way to test for poor HDR because we only screen after both transfection and HDR, but if you have good cutting and transfection efficiency and still no It is worth trying to pretreat with DMSO or a stimulus that opens the chromatin at your locus or to plate into media with M3814.It may also be worth trying to move to tag the other terminus if that is an option for your gene, as some termini just seem hard to tag for reasons we do not understand.You could also grow up some of the relatively brighter single-cell clones, amplify the locus using PCR, and analyze that sequencing, as it is possible that HDR was successful, but fluorescence intensity is just low.Some additional issues that may occur are described below.
Issue Suggested solution You only have heterozygous clones and want homozygous clones Try to screen more clones.In our experience, most knock-ins are either all homozygous or all heterozygous, but we have had one case (in a safe harbor locus) where there has been a mix that can be seen by screening more clones.

All the intermediate steps work and you still have no clones
There are a number of reasons this might be.While there are some genes that we have never been able to tag, here we include some options to try. 1) The chromatin at this locus may be very closed, and so pretreatment with something that would open the chromatin at that locus (DMSO or a specific inducer if the gene is inducible) may help.
2) Consider trying to tag the other terminus.3) The gene may be expressed too weakly to see fluorescence or the protein may be so dispersed in the cell that the fluorescence is not visible.In this case, one option is to make a transcriptional reporter by adding a T2A sequence between the endogenous protein and the fluorescent protein and adding an NLS to the fluorescent protein to concentrate the fluorescent protein signal in the nucleus.Another option is to use a brighter fluorescent protein or multiple copies of the fluorescent protein.You can generally assess if there is correct tagging but weak fluorescence by sorting the relatively brighter cells and screening those colonies by PCR to see if the DNA sequence is or is not inserted at the locus-if the correct DNA is there but you do not see fluorescence, it is likely too dim.Your tagged gene seems to have perturbed function Consider tagging the other terminus, adding a longer/different linker, or adding a T2A so it becomes a transcriptional reporter.You could also investigate smaller tags.

Software and datasets 1 . 1 )Figure 1 .
Figure 1.Diagram summarizing what will be accomplished in step A of this procedure.We will design multiple gRNAs, in this case targeting the C-terminal end of mouse IRF1, will clone each of them into a gRNA-Cas9 expression plasmid, and will test them to determine the optimal guide RNA (in this example, gRNA 1).The light pink or blue underlines in the sequence indicate the gRNA sequences, and the dark pink underlines indicate the PAM sequences for each possible gRNA.The desired insertion site is marked with yellow arrows, and the cleavage site for gRNA 1 is marked with purple arrows.

Figure 2 .
Figure 2. Example output from crispor.tefor.net(http://crispor.tefor.net/).Select the 100 bp region from 50 bp upstream to 50 bp downstream of your desired knock-in insertion position, enter it into cirspor.tefor.net,and then analyze the possible gRNA results to choose the three most promising guides based on having a high predicted efficiency, high specificity score, and being as close to your desired insertion site as possible.In this example, gRNAs 1, 2, and 3 best meet these criteria.

Published: Mar 20, 2024 b.
The next day (day 2), for each gRNA to test, transfect one well.i. Add 2.5 μg of pSpCas9-puro gRNA plasmid DNA to 125 μL of Opti-MEM in an Eppendorf tube.ii.Add 7.5 μL (for a 1:3 ratio with the DNA) of Lipofectamine 2000 transfection reagent to 125 μL of Opti-MEM in a second Eppendorf tube.iii.Add the Lipofectamine + Opti-MEM solution to the DNA + Opti-MEM solution, pipette up and down gently, and incubate for 5 min at room temperature.

Figure 4 .
Figure 4. Diagram of construction of homology-directed repair (HDR) donor plasmid.The plasmid

Figure 5 .
Figure 5. Graphical outline of steps C and D. We transfect the gRNA-Cas9 and homology-directed repair (HDR) donor plasmids into the cells, grow up transfected cells, sort single positive transfected cells into individual wells using FACS, and grow up clones for further screening.

4 .
Set up the Neon transfection system machine, program it for 1,680 V, 20 ms, 1 pulse, and set up the Neon cuvette with 3 mL of buffer E2 from the Neon electroporation reagent kit. 5. Prepare an Eppendorf tube with 1 mL of DPBS for washing the Neon tip. 6. Spin down 7.5 × 10 6 RAW 264.7 cells at 200× g for 3 min and aspirate the supernatant.

Figure 6 .
Figure 6.Sample field of view from imaging bulk transfected cells after they have grown to confluency where a gene has been tagged with YFP.In this field of view, there is a group of cells that express YFP after stimulus, which are the edited cells we are looking for.There are also many cells that do not express YFP.

15 Published: Mar 20, 2024 8 .
Cite as: Naigles, B. et al. (2024).Genetic Knock-ins of Endogenous Fluorescent Tags in RAW 264.7 Murine Macrophages Using CRISPR/Cas9 Genome Editing.Bio-protocol 14(6): e4960.DOI: 10.21769/BioProtoc.4960.Grow up single colonies into clonal populations.a. Seven to nine days after sorting, look at each plate under a regular tissue culture inverted microscope and identify wells with colonies growing in them.Note: We usually have between 30% and 60% of wells with growing colonies.b.Using a fluorescence microscope, image each growing colony in the channel where you expect to see your knock-in fluorescence.c.If you are looking for stimulus-responsive expression, add stimulus, wait the appropriate time for your gene to turn on, and then image on the fluorescence microscope again.d.Based on the fluorescence images, identify clones that are positive for fluorescence.We generally also

Figure 8 .
Figure 8. Sample images from screening clones in a 96-well plate eight days after sorting, showing both a positive clone and a negative clone for the knocked-in YFP tag in a nuclear marker (iRFP+) background

Figure 9 .
Figure 9. Cartoon example of a screening gel.Wild-type unedited cells have one small band, clones that do not have the knock-in inserted have one small band, homozygous knock-ins have one large band, and heterozygous knock-ins have one large band and one small band.

Figure 10 . 7 .
Figure10.Example of a Sanger sequencing chromatogram with these overlapping peaks.This example is from tagging CXCL10 (instead of the IRF1 example used for the rest of this protocol), as we had no homozygous knock-ins in IRF1 and never saw this overlapping peak phenomenon there.Based on this chromatogram, we chose to not move forward with this particular clone.Arrows point to specific regions of overlapping peaks.

3. Antibiotic-free DMEM Reagent Final concentration Quantity or Volume
Design guide RNAs a. Identify where in the mouse genome you want to insert your knock-in sequence.For endogenous gene tagging, this should either be immediately after the endogenous ATG or immediately before the endogenous stop codon.b.Select the genomic DNA sequence from 50 base pairs before to 50 base pairs after your desired insertion site.

Published: Mar 20, 2024 Component Volume (µL) pSpCas9
2.Clone the gRNA oligos into the pSpCas9-puro plasmid a.Anneal oligos.i.Dilute oligos to 100 µM each in autoclaved MilliQ water based on instructions Run the PCR product on a 1% agarose gel to confirm band size.d.Gel extract the PCR product from the gel using the QIAquick Gel Extraction kit.Send the uncut control and gRNA test amplicons that you gel extracted for Sanger sequencing with the primers you used for the PCR.
4. Extract genomic DNA from cell pellets a.We use the Zymo Quick-gDNA Miniprep kit to extract gDNA; whichever gDNA extraction kit you prefer should work.5.Amplify the region where the cut should occur a. Design PCR primers to amplify from ~300-600 bp upstream of the desired cut to ~300-600 bp downstream of the desired cut.Note: We use PrimerBlast (https://www.ncbi.nlm.nih.gov/tools/primer-blast/)from NCBI to identify primer pairs, aiming for 20 bp primers and an annealing temperature of 60 °C.b.Use these primers to amplify this region using PCR from the DNA you extracted from the gRNA test cells as well as the control uncut cells.Note: We use the NEB Phusion DNA polymerase and associated reagents, but alternative polymerase systems will likely also work.As the Phusion polymerase requires a different Tm for primers than is generally calculated, we use the NEB Tm calculator (https://tmcalculator.neb.com/#!/main) to calculate the Tm we will use in the PCR program.We use a thermocycler program of 98 °C for 30 s, then 30 cycles of (98 °C for 10 s, annealing temp for 30 s, 72 °C for 1 min per kilobase of PCR product), then 72 °C for 10 min, hold at 4 °C.c.Note: Any gel extraction kit should work.e.
You can use a different PCR chemistry, but we recommend a high-fidelity PCR enzyme because it is important for the entire plasmid to have the correct sequence.b.For N-terminal tagging, be sure to not include a stop codon on the fluorescent protein tag.c.We use PrimerBlast (https://www.ncbi.nlm.nih.gov/tools/primer-blast/) to design our cloning primers, aiming for 20 bp primers and an annealing temperature of 60 °C.d.We use restriction enzymes to digest 2 μg of the pUC19 backbone.We then run this digested backbone as well as our three PCR fragments (left homology arm, right homology arm, and fluorescent protein tag) on an agarose gel, gel extract the bands using the QIAquick gel extraction kit where we elute in 30 µL, and then run that product through a Zymo DNA Clean & Concentrator-5 kit and elute in 12 μL of the Zymo elution buffer.This two-step purification results in cleaner, higher-concentration DNA that leads to better efficiency with Gibson assembly in our hands.We do our Gibson assembly reactions using 100 ng of each DNA piece and use the NEBuilder HiFi DNA Assembly Master Mix, incubating for 1 h at 50 °C.We design our Gibson cloning to have ~23 bp overlaps between each two DNA pieces.e.If you are adding linker or T2A/P2A sequences into the plasmids between the endogenous protein and